Effects of past, present, and future ocean carbondioxide concentrations on the growth and survivalof larval shellfishStephanie C. Talmage and Christopher J. Gobler1

School of Marine and Atmospheric Sciences, Stony Brook University, Southampton, NY 11968

Edited by David M. Karl, University of Hawaii, Honolulu, HI, and approved August 31, 2010 (received for review December 3, 2009)

The combustion of fossil fuels has enriched levels of CO2 in theworld’s oceans and decreased ocean pH. Although the continuationof these processes may alter the growth, survival, and diversity ofmarine organisms that synthesize CaCO3 shells, the effects of oceanacidification since the dawn of the industrial revolution are not clear.Here we present experiments that examined the effects of theocean’s past, present, and future (21st and 22nd centuries) CO2 con-centrations on the growth, survival, and condition of larvae of twospecies of commercially and ecologically valuable bivalve shellfish(Mercenaria mercenaria and Argopecten irradians). Larvae grownunder near preindustrial CO2 concentrations (250 ppm) displayedsignificantly faster growth andmetamorphosis aswell as higher sur-vival and lipid accumulation rates compared with individuals rearedunder modern day CO2 levels. Bivalves grown under near preindus-trial CO2 levels displayed thicker, more robust shells than individualsgrown at present CO2 concentrations, whereas bivalves exposed toCO2 levels expected later this century had shells that were mal-formed and eroded. These results suggest that the ocean acidifica-tion that has occurred during the past two centuries may beinhibiting the development and survival of larval shellfish and con-tributing to global declines of some bivalve populations.

bivalve larvae | climate change | ocean acidification

More than 8 Pg of carbon dioxide (CO2) is released annuallyinto our planet’s atmosphere via the combustion of fossil fuels(1). About one-third of anthropogenically derived CO2 has enteredtheworld’s oceans during the past two centuries (2) and atmosphericand surface ocean CO2 levels are expected to reach ∼750 ppm by2100 (3, 4). CO2 entering the ocean decreases the availability ofcarbonate ions (CO3

−2) and reduces ocean pH, a process known asocean acidification. These changes in ocean chemistrymay have direconsequences for ocean animals that produce hard parts made fromcalcium carbonate (CaCO3). The experimental enrichment of CO2to levels expected in the coming century has been shown to dra-matically alter the growth, survival, and morphology of numerouscalcifying organisms including coccolithophores, coral reefs, crus-tose coralline algae, echinoderms, foraminifera, and pteropods (5–7). Many shellfish also produce calcareous shells, and juvenile andadult clams, mussels, and oysters have been shown to be adverselyaffected by elevated CO2 (8–12). The earliest life history stages ofshellfish, larvae, have been shown to be especially vulnerable to highCO2, displaying large declines in survival and delays in meta-morphosis at levels predicted to occur later this century, suggestingrecruitment of these populations may be adversely impacted byocean acidification (12–14).Although it is clear that calcifying ocean animals such as shellfish

are sensitive to the increases in CO2 projected for the future, theextent to which the rise in CO2 that has occurred since the dawn ofthe industrial revolution has impacted these populations is poorlyunderstood. Here we present experiments that examined theeffects of past (250 ppm), present (390 ppm), and future (>400ppm) CO2 concentrations on larvae of two species of shellfish: theNorthern quahog or hard clam,Mercenaria mercenaria, and the bayscallop, Argopecten irradians. These bivalves are ecologically and

commercially valuable resources: US mollusk harvests are $750million annually (15), with ecosystem services far exceeding thatvalue (16, 17). For experiments, CO2 was delivered via a gas pro-portionator system and CO2 levels in seawater were determined byquantifying dissolved inorganic carbon and pH during experimentsusing an EGM-4 Environmental Gas Analyzer (PP Systems) andthe program CO2SYS (http://cdiac.ornl.gov/ftp/co2sys/). Dissolvedinorganic carbon was measured with a methodological precisionof ±3.6% and full recovery (102 ± 3%) of Dr. Andrew Dickson’s(Scripps Institution of Oceanography, University of California atSan Diego, La Jolla, CA) certified reference material for total in-organic carbon in seawater [Batch 102 = 2,013 μmol dissolvedinorganic carbon (DIC) kg seawater−1] was obtained with our an-alytical procedures. Static delivery of CO2 at rates that turned overexperimental vessels several times an hour resulted in constant pHlevels during experiments [

Levels of CO2 strongly influenced the early formation ofM. mercenaria and A. irradians shells. For example, after 17 d ofdevelopmentM.mercenaria shells were 17± 2 μm thick under∼250ppm CO2, 6.7 ± 2 μm at ∼390 ppm CO2, and 3.8 ± 1 μm at ∼750ppm and ∼1,500 ppm CO2 (P < 0.001; Figs. 2 and 3). A. irradiansshells also decreased in thickness with increasing CO2 being 20± 3,12± 1, 11± 1, and 6.3± 1 μm thick under∼250, 390, 750, and 1,500ppm CO2 (P < 0.001, Fig. 2), respectively. Beyond impacting shellthickness, elevated levels of CO2 severely altered the developmentof the hinge structure of early stage bivalves. As CO2 levels in-creased from ∼250 to ∼1,500 ppm, there were dramatic declines inthe size, integrity, and connectedness of the hinge (Fig. 3). Al-though the M. mercenaria hinge displayed a “tongue and groove”pattern under low CO2 (250 and 390 ppm), under higher CO2

concentrations the hinge and associated hinge teeth became in-creasingly separated and detached. Given that the bivalve hingefacilitates opening and closing of shells, allowing for intake of foodand the excretion of waste (18), the compromised hinges observedunder elevated CO2 may hinder the ability of individuals to obtainand process suspended particles for nutrition. This hypothesis isconsistent with changes in lipid stores of larval shellfish exposed todiffering CO2 concentrations. For both species, with each in-creasing level of CO2, the lipid content (as estimated by an index)decreased significantly (P < 0.001; Fig. 2). Increasing CO2 con-centrations also caused marked changes in the morphology of theouter edge of juvenile shells (Figs. 3 and 4). With increasing levelsof CO2, this region of the shell became increasingly riddled withholes, pockmarks, and crevices, observations consistent with other

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Fig. 1. Development and survival of M. mercenaria and A. irradians larvae. Percent survival and developmental stage (veliger, pediveliger, and meta-morphosed) of larvae grown under four levels of CO2, ∼250, 390, 750, and 1,500 ppm (Table 1). The relative SD of larval survival among replicated vessels pertreatment for all times points and experiments was 4% (n = 4 per treatment).

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juvenile and larval shellfish reared under high CO2 (10, 14), sug-gesting CaCO3 shells were malforming and/or dissolving undermore acidic conditions. Altered shell morphology was also obviousin juvenile scallops that had distinct ridges, characteristic of laterstages of development, under preindustrial CO2, whereas individ-uals reared under higher CO2 conditions lacked ridges, a sign ofslower development (Fig. 4 and 19).Shell integrity is one of the most important lines of defense for

larval and juvenile bivalve shellfish, because shells provide physicalsupport for soft and delicate internal organs (20) and protectionfrombenthic and pelagic predators and suspended particles (21, 22).As such, the thinner, frailer shells displayed by early life historybivalves reared under modern day and elevated CO2 would likelymake individuals more vulnerable to predation and/or other envi-ronmental stressors. Similarly, within an ecosystem setting, larvaethat accumulate fewer lipids (Fig. 2) are generally slower to meta-morphose (19) and are more likely to perish once settled (23). Fi-nally, individuals with extended metamorphosis times (Fig. 1) andthat are smaller (Fig. 2) would be susceptible to greater rates ofpredation and natural mortality (23, 24). Hence, within an ecosys-tem setting, mortality rates of early life history bivalves that developunder modern day and higher CO2 levels would be expected to beeven greater than the rates observed during our experiments. Given

that bivalves in coastal areas naturally experience extremely highmortality rates in the transition from larvae to benthic juveniles (9),increases in mortality due to elevated CO2 could have profoundeffects on estuarine bivalve populations (5).Our findings regarding the effects of future CO2 levels on larval

shellfish are consistent with recent investigations of ocean acidifi-cation demonstrating that calcifying organisms will experiencedeclines in survival and growth, as well as malformed CaCO3 shellsand hard parts (25). However, our examination of the developmentof larval shellfish at levels of CO2 present before the indus-trialization of the planet provides important insight regarding thepotential effects ocean acidification has had on calcifying organismsduring the past two hundred years. Consistent with our findings,larval oysters (Crassostrea virginica) have displayed slightly largershell area when grown under preindustrial CO2 levels comparedwith modern levels (26).During the ∼24 million years before the industrial revolution,

atmospheric CO2 levels are estimated to have been relatively static,likely fluctuating in a narrow range significantly below the con-centrations present today (27, 28). Moreover, periods of higherCO2 before this era may not have been accompanied by lower pHand carbonate ion concentrations because the oceans may havebuffered the more gradual changes in CO2 that have occurredthrough geological history (3, 29). The evolution of calcification inocean animals is unknown, and the multiple forms of CaCO3 syn-thesized by modern day calcifiers (calcite, aragonite, amorphousCaCO3, and high magnesium CaCO3) differ widely in their vul-nerabilities to dissolution under lower pH (30). Although the pre-cise evolutionary tracks of modern bivalves remain somewhatuncertain (31), fossil evidence suggests that 906 of the 958 livinggenera of bivalve mollusks, including the species presented here,have a record that began in the mid- to late Cenozoic with thegreatest continuous increase in genera between ∼15 and ∼25 Mya(32), a period of estimated lower CO2 levels compared with today(27, 28). Together with our results, this suggests that ocean acidi-fication since the industrial revolution may have applied selectionpressure on modern marine bivalves and may continue to do so inthe future.The shallow marine environments that many marine bivalves

occupy can harbor dynamic levels of pH and CO2 (33, 34) and theprecise degree of phenotypic plasticity of survival among bivalvelarvae in the face of higher CO2 has not been established. Adap-tation and evolution could promote the proliferation of bivalvestrains that are more resistant to the increases in ocean CO2expected in the coming century and some calcifying organisms mayeven benefit from higher CO2 levels (25, 35). Importantly, however,the current rates of increase in atmospheric CO2 are significantlyfaster than any recorded in tens of millions of years (27, 28), sug-gesting this evolutionary challenge may be without precedent forextant calcifying species.A comparison of our two study species may provide insight into

future evolutionary pressure of ocean acidification on marine cal-cifiers. Globally, M. mercenaria has a larger, more diverse geo-graphic distribution (36) than A. irradians (37), an attribute thatgenerally provides resistance to evolutionary pressures (38) such asincreasing CO2 levels. In addition, predicted extinction rates arehigher for the marine mollusk family Pectinidae, which includesA. irradians, than the Veneridae family, which includes M. merce-naria (39). This information, combined with the more dramaticdeclines in survival displayed byA. irradiansunder higherCO2 levelscomparedwithM.mercenaria (Fig. 1), suggestsA. irradiansmay facea greater evolutionary challenge in adapting to future increases inCO2 concentrations.Precipitous declines in wild populations of bivalves during the

20th century have been attributed to overfishing, loss of habitat,hypoxia, and harmful algal blooms (40, 41). Our results suggestthat ocean acidification is another process that may have con-tributed to the declines of these populations in the recent past

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Fig. 2. Diameters, shell thickness, and lipid index of bivalve larvae grownunder a range of CO2 concentrations. Data are from four levels of CO2, ∼250,390, 750, and 1,500 ppm. (A) Diameters of M. mercenaria (day 24) and A.irradians (day 20). (B) Thickness of M. mercenaria (day 36) and A. irradians(day 52) shells at midpoint between the hinge and valve edge of the upperand lower shell of cross sectioned individuals. (C) Lipid index (lipid area/totalarea) for M. mercenaria (day 24) and A. irradians (day 20). Error bars rep-resent SD of replicated vessels per treatment (n = 4 per treatment).

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and could further impact bivalve population densities and di-versity in the future. Looking forward, marine organisms will bethreatened by aspects of climate change beyond elevated CO2,including higher temperatures. Given that the rise in oceantemperatures projected for the coming century (4) is withina range that could also hinder the growth and survival of bivalvelarvae (19, 42), future studies should consider the impact ofhigher CO2 in conjunction with temperature changes in line withsuch projections.

Materials and MethodsCO2 Treatments and Measurements. A gas proportionator system (Cole ParmerFlowmeter system, multitube frame) was used to deliver CO2 gas to seawatertreatments at multiple rates. The gas proportionator mixed appropriateflow rates of 5% carbon dioxide gas, low carbon dioxide gas, and pressur-ized air (∼390 ppm CO2) to yield the concentrations of carbon dioxide de-sired for experiments at a net flow rate (350 ± 5 mL min−1) that turned overthe volume of plexiglass covered experimental beakers >400 times daily.Experiments were repeated with tanked gas premixed at each specific CO2level and nearly identical seawater chemistry and larval responses wereobtained. For experiments, the CO2 gas mixtures from the proportionatorsystem were continuously delivered to the bottom of four replicated, poly-propylene 1-L beakers containing 0.2 μm filtered seawater from easternShinnecock Bay, NY. With continuous bubbling, all treatment beakersremained saturated with respect to oxygen (∼8 mg L−1). To quantify preciseCO2 levels attained in experimental beakers, seawater in beakers was bub-bled for 24 h and analyzed at the start (immediately before the addition oflarvae and phytoplankton) and at the end (larvae removed, phytoplanktonpresent) of each experiment using an EGM-4 Environmental Gas Analyzer(PP Systems) system that quantifies total dissolved inorganic carbon levelsafter separating the gas phase from seawater using a Liqui-Cel Membrane(Membrana). This instrument provided a methodological precision ±3.6%

for replicated measurements of total dissolved inorganic carbon and pro-vided full recovery (102 ± 3%) of Dr. Andrew Dickson’s (Scripps Institution ofOceanography, University of California at San Diego, La Jolla, CA) certifiedreference material for total inorganic carbon in seawater (batch 102 = 2,013μmol DIC kg seawater−1). Levels of CO2 were subsequently calculated basedon measured levels of total inorganic carbon, pH (total scale; mol kg sea-water−1), temperature (∼24 °C), salinity (∼28 ppt), and first and second dis-sociation constants of carbonic acid in seawater according to Roy et al. (43)using the program CO2SYS (http://cdiac.ornl.gov/ftp/co2sys/). Multiple dailymeasurements of pH (calibrated prior each use with NIST traceable stand-ards, ± 0.002, Orion Star Series Benchtop pH meter; Thermo Scientific) in-dicated experiment beakers maintained a constant pH level throughout allexperiments (

or detergents (45). To discourage the growth of bacteria during experi-ments, an antibiotic solution (5,000 units of penicillin, 5 mg of streptomycin,and 10 mg of neomycin per mL of solution, No.4083; Sigma-Aldrich) wasadded to each beaker at 1% its original concentration at the beginning of

each experiment and during each water change (approximately two timesweekly). This antibiotic mixture at this concentration has been shown tohave no negative effects on the growth and survivorship of shellfish larvae(45). For each experiment, ∼200 larvae were distributed to each experi-mental beaker, achieving an environmentally realistic abundance of larvae(42). Each treatment began with ∼900 mL to allow enough beaker volumefor the algal culture to be added daily as a food source. Twice weekly duringexperiments, larvae were gently poured onto a 64-μm mesh, and the con-dition (live or dead) and developmental stage of each larvae (veligers,pediveligers, and metamorphosed) was determined visually under a dissect-ing microscope; every individual larvae was counted at every water change.Larvae from each beaker (n = 4, per treatment) were removed, counted,observed, and transferred into a new beaker with new filtered seawater,food, and antibiotics within a 15 min period. Throughout experiments, allbeakers were submerged in a water bath maintained at 24 °C via the use ofcommercially available heaters and chillers. This temperature generallyyields high growth rates for A. irradians and M. mercenaria larvae (19, 42).Percent survivorship of all larvae was determined at each of the biweeklywater changes when the numbers of larvae in each stage of veligers, ped-iveligers, and metamorphosed juveniles were quantified. Experiments wereterminated after all surviving larvae in all treatments had metamorphosed.To statistically evaluate the effect of CO2 treatments on larval survival,goodness of fit tests (G Tests) were performed (46).

SEM. To document differences in the size and structure of larval and earlyjuvenile shellfish exposed to differing levels of CO2, randomly chosen indi-viduals (n = 4 per treatment) were mounted for SEM in two distinct ways.Firstly, to image the outside of shells, individuals were attached at 45° rel-ative to a level surface to a conductive substrate using carbon, double-sidedtape and were subsequently coated with ∼12 nm of gold using an Edwards150B rotary pump. To image the thickness and internal dimensions, cross-sections of shellfish were prepared. Individuals were mounted on glass mi-croscope slides using UV-curing adhesive coating (Locite 4304) and wereimpregnated with low-viscosity epoxy (Stuers’ Specifix-20) under vacuumoutgassing, a step that did not alter the original shape or size of individuals.After curing, the epoxy mount was progressively ground and polished to thecenterline (hinge to shell edge) of the shellfish using silicon carbide sand-papers, followed by successively finer diamond polishing grits (15, 6, and 3μm), 0.05 μm aluminum oxide suspension, and finally with colloidal silica. Allindividuals were cross-sectioned at the same location (hinge to shell edge)across the shell. This mount was then attached to a conductive substrateusing carbon double-sided tape and coated with ∼4 nm of gold. SEM imageswere collected on both types of samples with a Leo (Zeiss) Model #1550electron microscope using a high voltage of 20 KV and a Robinson back-scatter detector. All components of individual bivalve shells displayed in Figs.3 and 4 were probed using advanced EDAX/EDA microanalysis in the LEO

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Fig. 4. SEM images of 52-d-old A. irradians grown under different levels ofCO2:, ∼250, 390, 750, and 1,500 ppm, (Table 1). (A) Image of a full individuallarvae under each CO2 level. (B) A magnification of the outermost shell ofindividuals under each CO2 level.

Table 1. Temperature, pH, carbonate chemistry, alkalinity, and salinity (±SD) during the four-level CO2experiments with M. mercenaria, and A. irradians larvae

Parameter Near preindustrial CO2 Ambient, present day CO2 Year 2100 CO2 Year 2200 CO2

M. mercenariaTemperature (°C) 24 ± 0.52 24 ± 0.52 24 ± 0.52 24 ± 0.52pH 8.171 ± 0.022 8.052 ± 0.036 7.801 ± 0.004 7.532 ± 0.021pCO2 (ppm) 247.1 ± 6.231 380.0 ± 33.02 742.3 ± 9.111 1516 ± 31.21Ωcalcite 5.31 ± 0.47 4.53 ± 0.41 2.82 ± 0.05 1.67 ± 0.05Ωaragonite 3.42 ± 0.30 2.92 ± 0.26 1.82 ± 0.03 1.08 ± 0.03Total DIC (μmol L1) 1646 ± 94.21 1831 ± 52.34 1947 ± 21.33 2108 ± 18.06CO3

2− (μmol L−1) 208.0 ± 20.22 178.0 ± 16.03 111.0 ± 1.806 66.0 ± 1.904Alkalinity (TA) 1938 ± 117.3 2070 ± 66.42 2080 ± 22.63 2127 ± 49.71Salinity 28.0 ± 1.0 28.0 ± 1.0 28.0 ± 1.0 28.0 ± 1.0

A. irradiansTemperature (°C) 24 ± 0.51 24 ± 0.52 24 ± 0.52 24 ± 0.52pH 8.170 ± 0.026 8.041 ± 0.044 7.801 ± 0.005 7.530 ± 0.011pCO2 (ppm) 244.1 ± 4.006 386.5 ± 40.04 738.9 ± 9.941 1529 ± 35.05Ωcalcite 5.18 ± 0.06 4.55 ± 0.47 2.81 ± 0.06 1.66 ± 0.05Ωaragonite 3.34 ± 0.35 2.94 ± 0.30 1.81 ± 0.04 1.07 ± 0.03Total DIC (umol L−1) 1613 ± 53.54 1850 ± 30.98 1941 ± 25.54 2101 ± 9.221CO3

2− (μmol L1) 202.0 ± 23.42 180.0 ± 18.44 111.0 ± 2.341 66.02 ± 1.911Alkalinity (TA) 1899 ± 35.24 2090 ± 50.01 2075 ± 26.84 2146 ± 11.21Salinity 28.0 ± 1.0 28.0 ± 1.0 28.0 ± 1.0 28.0 ± 1.0

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(Zeiss) Model #1550 electron microscope and were confirmed to containalmost exclusively C, O, and Ca.

Size and Lipid Analysis. To estimate the relative lipid content of larvae, NileRed stain was used to bind to neutral lipids and fluoresce under an FITC filteron an epifluorescent microscope (23, 47). A Nile Red stock solution was madeof 1.25 mg of Nile Red crystals in 100 mL of acetone. Randomly selectedlarvae (n = 15) from each treatment were stained with a 1:9 dilution of thestock solution and 0.2 μm filtered seawater. Larvae were exposed to thestain for ∼1.5 h, rinsed with filtered seawater, and digitally photographedwith a Roper Scientific Photometrics CoolSNAP ES camera under an epi-fluorescent microscope. Digital images of each larva were analyzed for the

area of lipid accumulation and the diameter and the area of individualsusing ImageJ. A lipid index was estimated by dividing the area of the larvaecontaining the fluorescing lipids by the total larval area, thereby allowingfor direct comparisons among treatments. One-way ANOVAs and posthocTukey multiple comparison tests were performed to examine the differencesamong larval lipid indexes, shell length, and thickness, at each CO2 level.

ACKNOWLEDGMENTS. We are grateful for our supply of larvae from theEast Hampton Shellfish Hatchery. We thank Jim Quinn for SEM assistanceand James Waldvogel for cross sectioning assistance during this project.Constructive reviews came from two anonymous reviewers. This researchwas supported by the New Tamarind Foundation.

1. Le Quere C, et al. (2009) Trends in the sources and sinks of carbon dioxide. Nat Geosci2:831–836.

2. Sabine CL, et al. (2004) The oceanic sink for anthropogenic CO2. Science 305:367–371.3. Caldeira K, Wickett ME (2003) Oceanography: Anthropogenic carbon and ocean pH.

Nature 425:365.4. I.P.C.C. (2007) Intergovernmental Panel on Climate Change. Summary for Policy-

makers. The Physical Sciences Basis. Working Group I Contribution to the Fourth As-sessment Report of the IPCC, eds Solomon, et al. (Cambridge University Press,Cambridge).

5. Guinotte JM, Fabry VJ (2008) Ocean acidification and its potential effects on marineecosystems. Ann N Y Acad Sci 1134:320–342.

6. Kleypas JA, et al. (2006) Impacts of ocean acidification on coral reefs and other marinecalcifiers. Impacts of Ocean Acidification on Coral Reefs and Other Marine Calcifiers: AGuide for Future Research, Report of a Workshop Held 18–20 April 2005, St. Petersburg,Florida, Sponsored by NSF, NOAA, and U.S. Geological Survey (NOAA / Pacific MarineEnvironmental Laboratory, Seattle). Contribution 2897.

7. Riebesell U, et al. (2000) Reduced calcification of marine plankton in response toincreased atmospheric CO2. Nature 407:364–367.

8. Gazeau F, et al. (2007) Impact of elevated CO2 on shellfish calcification. Geophys ResLett 34:L07603:1–5.

9. Green MA, Jones ME, Boudreau CL, Moore RL, Westman BA (2004) Dissolutionmortality of juvenile bivalves in coastal marine deposits. Limnol Oceanogr 49:727–734.

10. Green MA, Waldbusser GG, Reilly SL, Emerson K, O’Donnell S (2009) Death bydissolution: Sediment saturation state as a mortality factor for juvenile bivalves.Limnol Oceanogr 54:1037–1047.

11. Kurihara H, Asai T, Kato S, Ishimatsu A (2008) Effects of elevated pCO2 on earlydevelopment in the mussel Mytilus galloprovincialis. Aquat Biol 4:225–233.

12. Kurihara H, Kato S, Ishimatsu A (2007) Effects of increased seawater pCO2 on earlydevelopment of the oyster Crassostrea gigas. Aquat Biol 1:91–98.

13. Talmage SC, Gobler CJ (2009) The effects of elevated carbon dioxide concentrationson the metamorphosis, size, and survival of larval hard clams (Mercenariamercenaria), bay scallops (Argopecten irradians), and Eastern oysters (Crassostreavirginica). Limnol Oceanogr 54:2072–2080.

14. Watson SA, Southgate PC, Tyler PA, Peck LS (2009) Early larval development of thesydney rock oyster Saccostrea glomerata under near-future predictions of CO2 drivenocean acidification. J Shellfish Res 28:431–437.

15. Cooley SR, Doney SC (2009) Anticipating ocean acidification’s economic consequencesfor commercial fisheries. Environ Res Lett 4:1–8.

16. Costanza R, et al. (1997) The value of the world’s ecosystem services and naturalcapital. Nature 387:253–260.

17. Cooley SR, Kite-Powell HL, Doney SC (2009) Ocean acidification’s potential to alterglobal marine ecosystem services. Oceanography (Wash DC) 22:172–180.

18. Eble AE (2001) Anatomy and Histology of Mercenaria mercenaria. Biology of the HardClam, eds Kraeuter JN, Castagna M (Elsevier, Amsterdam), pp 117–216.

19. Cragg SM (2006) Development, Physiology, Behaviour, and Ecology of Scallop Larvae.Scallops: Biology, Ecology, and Aquaculture, eds Shumway SE, Parsons GJ (Elsevier,Amsterdam), pp 45–122.

20. Carriker MR (1986) Influence of suspended particles on biology of oyster larvae inestuaries. Am Malacol Bull Special Ed 3:41–49.

21. Carriker MR (1996) The shell and ligament. The Eastern Oyster: Crassostrea virginica,eds Kennedy VS, Newwell RIE, Eble AE (Maryland Sea Grant College, University ofMaryland System, College Park, MD), pp 75–168.

22. Purcell JE, Cresswell FP, Cargo DG, Kennedy VS (1991) Differential ingestion anddigestion of bivalve larvae by the scyphozoan Chrysaora quinquecirrha and thectenophore Mnemiopsis leidyi. Biol Bull 180:103–111.

23. Phillips NE (2002) Effects of nutrition-mediated larval condition on juvenileperformance in a marine mussel. Ecology 83:2562–2574.

24. Tamburri MN, Zimmer-Faust RK (1996) Suspension feeding: Basic mechanismscontrolling recognition and ingestion of larvae. Limnol Oceanogr 41:1188–1197.

25. Doney SC, Fabry VJ, Feely RA, Kleypas JA (2009) Ocean acidification: The other CO2problem. Annu Rev Mater Sci 1:169–192.

26. Miller AW, Reynolds AC, Sobrino C, Riedel GF (2009) Shellfish face uncertain future inhigh CO2 world: Influence of acidification on oyster larvae and growth in estuaries.Plos ONE 4:e5661.

27. Pagani M, Zachos JC, Freeman KH, Tipple B, Bohaty S (2005) Marked decline inatmospheric carbon dioxide concentrations during the Paleogene. Science 309:600–603.

28. Pearson PN, Palmer MR (2000) Atmospheric carbon dioxide concentrations over thepast 60 million years. Nature 406:695–699.

29. Caldeira K, Berner R, Sundquist ET, Pearson PN, Palmer MR (1999) Seawater pH andatmospheric carbon dioxide. Science 286:2043a.

30. Mann S (2001) Biomineralization: Principles and Concepts in Bioinorganic MaterialsChemistry (Oxford University Press, New York).

31. Harper EM, Taylor JD (2000) The Evolutionary Biology of the Bivalvia, ed JAC,(Geological Society, London, Special Publications,).

32. Jablonski D, Roy K, Valentine JW, Price RM, Anderson PS (2003) The impact of the pullof the recent on the history of marine diversity. Science 300:1133–1135.

33. Feely RA, Sabine CL, Hernandez-Ayon JM, Ianson D, Hales B (2008) Evidence forupwelling of corrosive “acidified” water onto the continental shelf. Science 320:1490–1492.

34. Salisbury J, Green M, Hunt C, Campbell J (2008) Coastal acidification by rivers: A newthreat to shellfish? Eos Trans AGU 89:513.

35. Ries JB, Cohen AL, McCorkle DC (2009) Marine calcifiers exhibit mixed responses toCO2 induced ocean acidification. Geology 37:1131–1134.

36. Harte ME (2001) Systematics and Taxonomy. Biology of the Hard Clam, edsKraeuter JN, Castagna M (Elsevier, Amsterdam), pp 3–43.

37. Brand AR (2006) Scallop ecology: Distributions and behaviour. Scallops: Biology,Ecology, and Aquaculture, eds Shumway SE, Parsons GJ (Elsevier, Amsterdam), pp651–713.

38. Jablonski D (1987) Heritability at the species level: Analysis of geographic ranges ofcretaceous mollusks. Science 238:360–363.

39. Roy K, Hunt G, Jablonski D (2009) Phylogenetic conservatism of extinctions in marinebivalves. Science 325:733–737.

40. Gobler CJ, Lonsdale DJ, Boyer GL (2005) A review of the causes, effects, and potentialmanagement of harmful brown tide blooms caused by Aureococcus anophagefferens(Hargraves et Sieburth). Estuaries 28:726–749.

41. Jackson JBC, et al. (2001) Historical overfishing and the recent collapse of coastalecosystems. Science 293:629–637.

42. Carriker MR (2001) Embryogenesis and Organogenesis of Veligers and Early Juveniles.Biology of the Hard Clam, eds Kraeuter JN, Castagna M (Elsevier, Amsterdam), pp77–115.

43. Roy RN, et al. (1993) The dissociation constants of carbonic acid in seawater atsalinities 5 to 45 and temperatures 0 to 45 °C. Mar Chem 44:249–267.

44. Riebesell U, Fabry VJ, Hansson L, Gattuso JP (2010) Guide to Best Practices for OceanAcidification Research and Data Reporting (Publications Office of the EuropeanUnion, Luxembourg).

45. Zeebe RE, Zachos JC, Caldeira K, Tyrrell T (2008) Oceans. Carbon emissions andacidification. Science 321:51–52.

46. Padilla DK, Doall MH, Gobler CJ, Hartson A, O’Boyle K (2006) Brown tide alga,Aureococcus anophagefferens, can affect growth but not survivorship of Mercenariamercenaria larvae. Harmful Algae 5:736–748.

47. Sokal RR, Rohlf FJ (1995) Biometry: The Principles and Practice of Statistics inBiological Research (W.H. Freeman and Company, New York), 3rd Ed.

48. Castell LL, Mann R (1994) Optimal staining of lipids in bivalve larvae with Nile Red.Aquaculture 119:89–100.

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